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Composition analysis of gastrointestinal contents of laboratory‐reared and wild lake sturgeon (Acipenser fulvescens)

Composition analysis of gastrointestinal contents of laboratory‐reared and wild lake sturgeon... INTRODUCTIONConservation history of Acipenser fulvescensSturgeon belong to the family Acipenseridae, one of the most widespread families of fishes worldwide, inhabiting estuaries, rivers, near‐shore oceanic environments and inland seas of the Northern hemisphere (Pikitch et al., 2005). There are 25 extant Acipenserid species (Shen et al., 2020), including the species this study focuses on, Acipenser fulvescens (Rafinesque, 1817). This species evolved in the Jurassic period 250 million years ago and is considered a “living fossil” (Birstein, 1993; Harkness & Daymond, 1961) but suffered substantial population declines due to anthropogenic activities like habitat degradation, habitat fragmentation, and overharvesting that were common at the start of the 20th century (Pollock et al., 2015). They are primarily found in the Great Lakes and Mississippi River drainage (Ferguson & Duckworth, 1997), but they are also native to the Coosa River basin which crosses north central Georgia, southeastern Tennessee and northeast Alabama.Around 1960, the original population of A. fulvescens in the Coosa, Tennessee and Cumberland Rivers became extirpated by overharvesting due to increased demand for caviar and degraded water quality related to urbanization and intensive dam construction throughout the southeastern United States (Walker, 2017). To restore A. fulvescens to those rivers, Georgia Department of Natural Resources Wildlife Resources Division (GA‐DNR) initiated a long‐term stocking programme in the Coosa and Tennessee Rivers with the goal of re‐establishing the population of this native sport fish to Georgia and address the conservation of Georgia's rare species. Juvenile A. fulvescens were first stocked by GA‐DNR in the Coosa River system from 2002 to 2005, releasing 45,091 fingerlings during this period (Bezold & Peterson, 2008). After 4 years, an assessment study showed an increase in abundance of juveniles to an overall population estimate of n = 789 (690–889, 95% confidence interval). Each cohort's survival rate ranged from 1% to 14%, depending on the size of fish stock in each year (Bezold & Peterson, 2008), and the survival rate through 2019 is estimated to be 1% overall for the Coosa River, and 2.5% for the Tennessee River. Recovery is expected to be slow due to the lengthy juvenile stage. Sexual maturity as indicated by first spawning occurs relatively late, typically at age 12–15 for males (Bruch, 1999; Bruch & Binkowski, 2002; Peterson et al., 2007), and even later for females. One advantage for stocking success is that growth rate is faster in juveniles (Peterson et al., 2007). A similar restoration programme in Lake Oneida, New York resulted in a growth rate of 145 mm per year for A. fulvescens aged 1–5 years (Jackson et al., 2002). Despite the slow rate of maturation and long‐life span of this species, this evidence suggests that traditional stocking programmes can be utilized to support and re‐establish self‐sustaining A. fulvescens populations. The ongoing efforts in the southeastern United States have been mostly successful reintroducing a stable population, but the aspect with greatest scope for improvement is in increasing stocking survival by reducing the impact of stocking stressors such as transitions in feed.Feeding of Acipenser fulvescensFeeding by A. fulvescens is characterized as generalist and opportunistic, with larvae subsisting on zooplankton (Muir et al., 2000), and after that they consume a wide variety of benthic organisms (Klassen & Peake, 2007; Nilo et al., 2006). The feeding habits of A. fulvescens vary seasonally, and it also depends on the availability of different foods in different seasons (Barth et al., 2013). Their primary foods include amphipods, oligochaetes, ephemeropteran nymphs, trichopteran larvae, mollusks and fish eggs (Nilo et al., 2006). In the Tennessee River, juveniles have been shown to feed selectively on larval Chironomids which vary seasonally in availability (Amacker & Alford, 2017).In the hatchery, chopped or whole bloodworms (Chironomus larvae), brine shrimp (Artemia) nauplii or other raw foods are commonly fed to A. fulvescens at the larval and adult stages (Table 1) (Aloisi et al., 2006; Klassen & Peake, 2007, 2008). Fishes fed with live feeds typically have a higher growth rate and lower mortality rate than those provided formulated food (Bauman et al., 2016). Still, live foods can have many disadvantages like high cost, irregular availability, and uncertainty about nutritional quality. According to Dvorak (2009), there is a risk of transmitting an unknown disease from hatchery‐reared fish to the environment if live feeds are fed to fishes. Formulated feed is generally more cost‐effective, is easier to manage and can be developed with stable and enriched nutrients (Hamre et al., 2013). In one study, an experimental diet with 3%–6% krill or fish protein content significantly improved growth of juvenile A. fulvescens and furthermore indicated that stimulation of feeding activity in this species shifted over the course of juvenile growth (from 2 to 280 g) (Moreau & Dabrowski, 1996). Juvenile A. fulvescens can be fed formulated diets (soft‐moist or dry) without impairing their survival and growth performance (Lee et al., 2018), but the formulated diets available for sturgeon almost certainly differ in their nutritional value from the wild diet (Table 1). At stocking, the transitions from one diet in captivity to a different food source in the wild may create additional stocking stress and act as a barrier to high post‐stocking survival rates of captively reared juveniles. Therefore, it is crucial to know about the composition of both captive and wild diets of juvenile A. fulvescens to gain an understanding of nutritional impacts occurring at stocking in order to optimize programmes that aim to help restore severely depleted or extirpated populations.1TABLEComposition of provided laboratory feeds per the supplier's guaranteed analysisDietSoft‐moist pelleted feedaFrozen krillbFrozen bloodwormsbProtein (%)45.06.53.5Fat (%)19.03.40.3Fibre (%)<2.00.70.7Ash (%)<9.01.31.6Moisture (%)25.088.093.5Note: Bloodworms approximate primary foodstuffs consumed by lake sturgeon in the Tennessee river (Amacker & Alford, 2017).aRangen Brand.bSan Francisco Bay Brand.Study objectivesGrowth of fish and feed utilization are influenced by the balanced nutritional composition of fish feed. An adequate supply of nutrients, both in terms of quantity and quality, should be met for good growth, survival and reproduction of fish and other aquatic animals (Hasan, 2001). During the transition from endogenous to exogenous feeding and for several weeks after the onset of exogenous feeding, larval fish in aquaculture typically exhibit periods of low growth and high mortality (Kamali et al., 2006). Similarly, A. fulvescens experience a shift in diet composition and availability during stocking to the river, and this may be associated with reduced survival. Therefore, it is important to evaluate and assess the proximate composition of food from both laboratory and wild habitats to allow for comparison of the nutritional value in these two systems and estimate the scale of the impact this dietary shift has post‐stocking. It is also required to know about the physiological system, dietary preference, digestive physiology and gastrointestinal anatomy to develop a successful feeding programme for A. fulvescens (Hofer et al., 1982). This study assessed the nutrient uptake from the diet of laboratory‐reared and wild A. fulvescens and how the hatchery rearing environment and riverine habitat differ from each other in available nutrition. In the present study, the stomach and colonic contents of wild A. fulvescens needed to be collected to assess the proximate composition of the diet. However, it is not feasible to sacrifice A. fulvescens to know the content of this fish's food due to its reduced population status. A safe and efficient technique was also needed to ensure a quick recovery by the wild‐caught fish after collecting contents from the gastrointestinal tract. Therefore, established techniques for digesta collection were first validated using controlled laboratory trials and then applied to samples collected from wild fish. The collected samples were then analysed for proximate composition in order to provide insight to nutrient processing in hatchery‐raised A. fulvescens fed a formulated diet, compared to juveniles subsisting on the natural diet available in the wild. We first validated gastric lavage and colonic flushing in the lab to support the collection of stomach and faecal contents from wild fish and then compared the proximate composition of stomach and colonic contents between the laboratory and wild environment. This comparison will support the conservation of this protected species, helping to re‐establish the population throughout its endemic range.MATERIALS AND METHODSExperimental animalsFour 150‐gal freshwater tanks were set up with water quality and physical conditions according to standard parameters for A. fulvescens (21°C, pH 7.18 ± 0.02, conductivity 470 ± 3.21 μS, dissolved oxygen held at 100% air saturation by continuous aeration). Nineteen 2.5 year‐old A. fulvescens individuals, ranging from 0.39 to 0.56 m total length (Figure 1) and weighing 0.34 ± 0.04 kg, were received from the US Fish & Wildlife Service Warm Springs National Fish Hatchery, brought to the University of West Georgia and evenly distributed among the prepared tanks. Fish were fed 2.4 mm diameter Rangen pellets and krill (Table 1) to satiation daily at 8 AM, following the standard diet and feeding procedures of the rearing hatchery. Leftover food was siphoned out after feeding activity stopped (approximately 45 min) to maintain water quality. Fish were acclimated to laboratory conditions for 1 month prior to the start of experiments, and water quality was monitored by measuring the concentrations of nitrate, nitrite, ammonia, pH, hardness and conductivity every day for the first 15 days after introduction of fish into the laboratory environment, and after that water quality parameters were checked once every 3 days, with a daily water exchange of 20–30 gal of water in each tank.1FIGUREComparison of total length (mm) of juvenile Acipenser fulvescens utilized for validation of gastric lavage and colonic flushing protocols versus wild A. fulvescens captured from the Tennessee River.Techniques for collection of gastrointestinal contentsGastric lavage is a non‐lethal method for examining fish stomach content from live fish, where stomach contents are flushed out with an apparatus inserted into the fish's pharynx. The gastric lavage technique has been tested in several sturgeon species. However, most of these studies have analysed the stomach contents for identifying prey items or determining mortality rates of the procedure itself, rather than determination of dietary nutritional profile (Haley, 1998; Wanner, 2006). Colonic flushing is a non‐lethal method for examining faeces in the intestinal tract. Deionized (DI) water is pulsed into the colon through the anus using a syringe attached with a smaller diameter tube. The tube is inserted gently into the colon and creates short pulses to expel all the faeces. This process continues until the expelled water is clear. Colonic flushing is less time‐consuming (15–30 s) and safer for A. fulvescens than the gastric lavage technique (Amacker, 2016).Laboratory validation of gastric lavage and colonic flushingAfter acclimatizing fish to the laboratory environment, we performed non‐lethal gastric lavage by following the protocol as described by Haley (1998). Before starting gastric lavage, total weight and total length of each fish were recorded, and then gastric lavage and colonic flushing were performed at six different time points (2, 4, 6, 12, 24 and 32 h after feeding) to determine the required digestion time, using 18 randomly selected fish (n = 3 fish per time point) in two trials. In the first trial, 12 fish were used to perform gastric lavage at 2, 4, 6 and 12 h post‐feeding. In the second trial, gastric lavage was done at 4 and 12 h post‐feeding on six fish, and the same fish were used for gastric lavage at 24 and 32 h post‐feeding after 1 week. Stomach content at 0 h post‐feeding was represented by analysis of the offered feed. Colonic flushing was not performed at 0 h post‐feeding as the colon was assumed to be empty at this time point. Fish were fasted the day before performing gastric lavage and fed Rangen and krill at the start of each trial. Table 1 shows the composition of these feed as given by the manufacturer's guaranteed analysis. Ingestion of feed by size‐matched fish was assumed to be evenly distributed within the tank. When the fish's feeding activity stopped, all leftover food was siphoned from the tank. The water was filtered with a very fine mesh net and the leftover food was weighed to determine the total feed ingested. For the gastric lavage protocol, each fish was anesthetized in a 5‐gal bucket containing 30 mL of tricaine methanesulfonate (MS‐222; 100 mg/L) buffered with sodium bicarbonate (200 mg/L) combined with 20 L of water, for a final immersion concentration of 0.15 mg/L MS‐222. Typically, fish were sedated after approximately 4 min. Once sedated, the fish was placed ventral side up at a 45° angle on a gastric lavage stand where flexible tubing attached to a 60‐cm3 syringe fitted with a 16‐ga needle was inserted into the stomach through the pharynx. A different diameter tube was used for different‐sized fish (Table 2), and the large diameter tubing omitted the needle fitting and was instead connected directly to the syringe. Pulsing DI water at the cardiac valve stimulated relaxation and allowed entry of the tube into the stomach. Water was pulsed by syringe into the stomach until it was firm. Then, a gentle massage of the ventral surface of the sturgeon was done to help release water and stomach contents into a pre‐weighed sample collection bottle. This procedure was continued until only clean water came out from the stomach. Haley (1998) suggested pulsing DI water for 3 min and repeated it three times for each animal. However, in this study, the timing to complete gastric lavage depended on fish size and the amount of food they had in their stomach but never exceeded 5 min. After gastric lavage, the fish was kept in a separate bucket filled with aerated fresh water until it recovered from the stress of the procedure and resumed regular activity. For most fish, some stomach content came out during the recovery period, and this was collected using a disposable pipette.2TABLEDiameter of gastric lavage tube according to fish size (Acipenser fulvescens)Fish total length (m)Outer diameter (OD) of lavage tube (mm)Inner diameter (ID) of lavage tube (mm)0.15–0.302.421.660.31–0.454.003.150.46–1.05.154.00Colonic flushing is a non‐lethal process to examine the faeces of the intestinal tract. This process used a 60‐cm3 syringe fitted with appropriate diameter tubing (Table 2) filled with DI water and inserted the tube end gently 30–50 mm into the colon through the anus. Faecal content was collected in a bottle using the same process as described above for gastric lavage and measured for total weight. Then, the solid and aqueous parts of both stomach and colonic contents were separated using a very fine mesh filter and centrifuging the samples. The weight of both aqueous and solid parts was measured, and the samples were subsequently stored at −20°C.Stomach and colonic contents collection from wild Acipenser fulvescensStomach and colonic contents were collected from A. fulvescens caught from the upper Tennessee River over 2 days in December 2019. Trotlines were used to catch the fish from the Watts Bar Reservoir in the drainage of this river (35.6206°N, 84.7832°W). The water temperature was 10.9°C, and air temperature was 4°C on the first day of sample collection and on the second day water temperature and air temperature were 10.3 and 2°C, respectively. Dissolved oxygen in the river was variable between sampling times, on average 10.38 ± 0.32 mg/L. Trotlines were set in the river with buffalo and carp baits at 4:30 PM and retrieved at 7 AM the following day (14.5 h total soak time). After getting the fish on the boat, the total and fork lengths and the total weight of each fish were measured. Once the trotlines were removed from the water, fish were transported in a live well to the riverbank where a 150‐gal tank was set up for keeping fish while collecting gut and colonic contents. Stomach and colonic contents of these different sized A. fulvescens (n = 15) were collected using the same procedures described before for laboratory fish and then released to the river. The size of tubing used for the gastric lavage was varied according to the fish's size, as given in Table 2. Wild fish had an average total length of 0.65 ± 0.74 m, ranging from 0.26 to 1.08 m (Figure 1). A tube of the same diameter (inner diameter 1.14 mm, outer diameter 1.57 mm) was used to collect colonic content for every fish size to provide more space for colonic content to come out. The bottles of the stomach and colonic content were transported on ice in a cooler to the University of West Georgia Aquatics Research Laboratory and stored at 4°C prior to filtration and processing and subsequently stored at −20°C for further analysis.Proximate composition analysis of stomach and colonic samplesAll stomach contents and faecal samples collected from wild and hatchery‐reared A. fulvescens were prepared and analysed according to the same protocols. The solid portion of the sample was separated by filtration using fine nylon mesh (approx. 0.3–0.5 mm pore size) followed by centrifugation at 10,000 × g for 10 min and stored at −20°C. Each solid sample was diluted by adding 2x (by weight) of the corresponding aqueous portion previously separated by filtration and then an electric tissue homogenizer was used for homogenization. Thus, both the solid and liquid fractions of the original sample were represented in the analysed sample. For the 0 h time point, thawed krill and Rangen pellets were separately diluted and homogenized. Heating of homogenate was prevented by using a short pulse (<5 s). The crude homogenate was separated into three aliquots for analysis of lipid, protein and carbohydrate. Aliquots for the water‐soluble protein and glucose fractions were centrifuged at 10,000 × g for 10 min to remove solid fragments, and the supernatant transferred to a new tube for analysis. The Folch method (Folch et al., 1957) was used for extraction of lipids from the remaining aliquot. In brief, 20× by volume (1500 μL) of 2:1 (v/v) chloroform:methanol solution was added to a 75 μL aliquot of the crude homogenate, mixed by vortexing and then kept on ice for 5 min. Then 0.2× (315 μL) 0.05% CaCl2 solution was added and centrifuged at 12,000 × g for 5 min. After centrifuging, the lower phase was separated to a new tube and used to measure the lipid content (mg/dL).Each processed sample aliquot was analysed for proximate composition using colorimetric assay kits, according to the manufacturer's protocols. Total protein (Sigma‐Aldrich, 96 Well Plate Bradford Assay Protocol, B6916), triglyceride (Cayman Chemical, No. 10010303) and glucose (Cayman Chemical, No. 10009582) were measured to give the concentration (mg/dL) of each of these macronutrients in each collected gut sample. In the Sigma protein assay protocol, a protein standard of bovine serum albumin (10–160 mg/dL) was used to make the standard curve, and 5 μL of each unknown sample was pipetted into other wells on the same plate. After initiating the reaction with Bradford reagent, the absorbance was measured at 595 nm at 25°C. For the lipid assay, 10 μL triglyceride standard of 0–200 mg/dL concentration was used for the standard curve, and 10 μL of each unknown sample was measured on a 96‐well plate, according to the kit manufacturer's instructions. The plate was incubated at 25°C for 15 min after adding 150 μL diluted enzyme buffer. Then the lipid concentration was measured by absorbance at 540 nm. For measurement of the glucose concentration, 15 μL of standard solution at 0–25 mg/dL concentrations established the standard curve. To each well, 85 μL of diluted NP‐40 assay buffer was added, and 100 μL of reconstituted enzyme mixture initiated the reaction. Absorbance was measured at 520 nm after incubation at 37°C for 10 min. Each unknown sample and standard solutions were measured in triplicate for all protocols.Statistical analysisAll data are presented as a mean ± SEM and were analysed using GraphPad Prism software (version 6.07). Multiple t‐tests were used to compare concentration of protein, lipid and glucose between the treatment groups. To compare the concentrations of protein, lipid and glucose among different time points, one‐way ANOVA was used, followed by Tukey's multiple comparisons post hoc test to examine pairwise comparisons. Values of p < 0.05 were considered significant, represented by an asterisk (*).RESULTSEfficiency of gastric lavage and colonic flushingGastric lavage and colonic flushing were used to collect stomach and colonic contents in both laboratory‐reared and wild A. fulvescens. The highest average percentage of ingested feed, 75.82% ± 14.09%, was recovered from the stomach at 4 h post‐feeding, with substantially lower amounts recovered at 2 h (36.67% ± 5.09%), 6 h (17.80 ± 4.01) and at 12 h (32.90% ± 17.93%) in the laboratory. No food was recovered after 12 h (Table 3). The amount of recovered colonic content increased with increasing time, and the amount per fish body mass (g/kg) was highest at 32 h post‐feeding (Table 3).3TABLEAmount of stomach and colonic content recovered from Acipenser fulvescens using gastric lavage and colonic flushing at 2–32 h post‐feeding in the laboratory (n = 3–6 per time point) as a percent of feed ingested (%), and relative to fish mass (g/kg)Post‐feeding time (h)Recovered stomach content (%)Stomach content (g/kg)Colonic content (g/kg)236.67 ± 5.090.29 ± 0.050.63 ± 0.29475.82 ± 14.090.96 ± 0.431.00 ± 0.40617.80 ± 4.010.18 ± 0.041.16 ± 0.401232.90 ± 17.930.32 ± 0.141.26 ± 0.3224003.01 ± 0.7932005.40 ± 1.30Average27.20 ± 11.620.29 ± 0.142.08 ± 0.75Note: Fish were fed a total of 0.33 g Rangen soft‐moist pellets and 2.33 g krill per fish.Following the protocol of gastric lavage and colonic flushing used in the laboratory, stomach and colonic contents were collected from A. fulvescens captured from the Tennessee River. We used a total of 15 fish for gastric lavage and colonic flushing in the wild habitat. One fish was not used for gastric lavage among those fish, and another fish was not used for colonic flushing because those fish were not in good enough condition to remain out of the water for the time required to perform both procedures. The amount of food ingested by A. fulvescens in the river was estimated based on the amount of recovered stomach solids, using digestion rates observed in the lab fish. In the lab, the average recovered stomach solid over 12 h was 1.23 g, and the amount of krill ingested per fish was 2.33 g (Table 3), indicating a recoverable proportion of 1.89, which was multiplied by the mean weight of stomach solids recovered from wild fish (2.72 g, Table 4) resulting in an estimated 5.15 g of food ingested by wild fish prior to capture. The average stomach solid recovery rate adjusted for fish mass (1.41 ± 0.51 g/kg) was lower than the colonic solid recovery rate (5.85 ± 0.94 g/kg) (Table 4). In wild fish, variation between individuals was greater, but on average, the amounts of solid content collected from the stomach and distal intestine were both roughly threefold greater compared to the A. fulvescens sampled in the lab (Table 3).4TABLEAmount of stomach and colonic content collected from Acipenser fulvescens (n = 15) by gastric lavage or colonic flushing at Tennessee RiverFish mass (kg)Stomach solid (g)Stomach solid recovered (g/kg)Colonic solid (g)Colonic solid recovered (g/kg)1.590.320.197.614.791.912.211.157.003.672.042.901.429.834.801.092.332.109.008.231.130.300.267.466.531.9513.807.08N/AN/A1.870.310.161.520.801.360.620.4416.9012.421.465.703.919.566.512.340.810.3417.127.281.630.530.318.124.901.630.800.499.655.881.891.440.7423.2112.295.266.211.1842.548.086.44N/AN/A9.931.54Average2.24 ± 0.392.73 ± 0.991.41 ± 0.5112.82 ± 2.695.85 ± 0.94Composition of stomach contents at different time pointsA total of 18 fish were used to evaluate gastric lavage efficiency in the laboratory at 2, 4, 6, 12, 24 and 32 h after feeding. The proximate composition was analysed individually at each sampling time. There was a significant difference in protein concentration in stomach (p = 0.0074, Figure 2A) over time but not in the colonic content (p = 0.3853, Figure 2B) over the six different time points examined in laboratory‐held A. fulvescens. Tukey's multiple comparison test identified that protein content was significantly lower than the provided feed (0 h) at every time point for which stomach contents were recovered (2, 4, 6 and 12 h). Lipid concentration (mg/dL) in recovered stomach and colonic contents was also assessed at the same post‐feeding time points. There was no significant difference in lipid concentration in stomach (p = 0.3174, Figure 2C) or colonic content (p = 0.1057, Figure 2D) among the measured time points in laboratory A. fulvescens. Glucose concentration in both stomach (p = 0.0240, Figure 2E) and colonic (p = 0.0480, Figure 2F) content was significantly different over the observed time frame. Interestingly, Tukey's multiple comparison test did not identify any significant differences among different time points in colonic glucose concentration (mg/dL), indicating that only the overall difference was significant for these samples. However, in stomach content, it showed a significant difference between 0 h compared to all subsequent time points (2, 4, 6 and 12 h).2FIGUREProtein, lipid and glucose concentration (mg/dL) in stomach (A, C, E) and colonic (B, D, F) contents from laboratory acclimated Acipenser fulvescens at 0–32 h post‐feeding. Stomach values at 0 h are the measured concentrations in the offered feed as determined by analysis identical to methods used for collected digesta. Colonic samples were not collected at 0 h. One‐way ANOVA indicated a significant difference in protein concentration in stomach content (p = 0.0074, A) but not in colonic content (p = 0.3853, B). Lipid concentrations (mg/dL) in both stomach (p = 0.3174, C) and colonic (p = 0.1057, D) contents at different post‐feeding time points were not significantly different. Glucose concentration (mg/dL) in stomach (p = 0.0240, E) and colonic (p = 0.0480, F) contents at different post‐feeding time‐points shows significant differences. Post hoc differences from 0 h, as determined by a pairwise Tukey's post hoc test, are indicated by “*”.Proximate composition of field‐collected stomach and colonic contentSeparated solid content from stomach and colon were used for proximate composition analysis by colorimetric assay protocol. Protein concentrations in stomach contents between laboratory fishes and wild fish were not significantly different (p = 0.2104) (Figure 3A), whereas there was a significant difference (p = 0.0187) in lipid concentration of stomach contents, with A. fulvescens fed the laboratory diet exhibiting substantially greater lipid content than wild‐collected fish. The stomach contents of fish from both habitats showed no difference (p = 0.6835) in glucose concentration (Figure 3A). A slightly different pattern was observed for colonic solids, with no differences between sampling groups in protein or lipid content, but glucose was significantly higher in wild fish (Figure 3B). Glucose was by far the smallest fraction contributing to the proximate composition of stomach and colonic contents of A. fulvescens in both the laboratory and wild habitats.3FIGUREComparison of protein, lipid and glucose concentrations (mg/dL) in the stomach solids (A) and colonic solids (B) of laboratory‐reared and wild Acipenser fulvescens. Asterisk symbol “*” indicates significant difference (p < 0.05) in concentrations between wild and laboratory‐reared fish as determined by multiple t‐tests, for protein (p = 0.2104), lipid (p = 0.0187) and glucose (p = 0.6835) of stomach contents (A) and protein (p = 0.6198), lipid (p = 0.1483) and glucose (p = 2.52e – 006) of faeces (B).DISCUSSIONFood transit time in alimentary canalThe amount of colonic content increased with time after feeding the fish in the laboratory. According to Venero et al. (2015), defecation is started between 8 and 16 h after feeding in the Gulf Sturgeon (Acipenser oxyrinchus desotoi), but in the present study faecal content was collected as early as 2 h after feeding. The fish were fasted for 24 h before each round of colonic sampling. As the total food transit time for A. fulvescens has been previously reported to be 32 h at 21°C (Venero et al., 2015), the faecal content collected at 2 h post‐feeding must have been from the day before colonic flushing was performed, that is, the residual of food ingested prior to the fasting period.The colonic flushing technique was efficient for all fish between 0.26 and 1.08 m, and larger amounts of faecal content on a per mass basis were collected from most of the wild fish than from the laboratory fish. Comparatively, the amount of faecal content was more than the amount of stomach content in wild A. fulvescens. The reason may be attributed to the amount of time the fish was on the hook of the trotline. The trotlines were in the river for approximately 14 h; if the fish was hooked at the beginning of the soak time, then it was not able to eat anything during this period. Thus, all the stomach contents will be digested, and the colonic content will increase, but as our laboratory experiments confirm, would likely still be present in the alimentary tract. Aside from consumption of the catch bait, the timing of the most recent feeding event for wild A. fulvescens cannot be known. The average amount of stomach solid recovered (1.41 ± 0.51 g/kg) from wild fish was higher than the laboratory fish (0.29 ± 0.14 g/kg); however, the weight‐specific collection rate for wild fish was quite similar to the laboratory‐based validation trial, indicating that recovered stomach contents were equally representative of the wild diet as in the laboratory validation trial, and was being digested at a comparable rate despite the 10°C difference between water temperatures in the laboratory and the river. As estimated by back calculating from the recovered amount of stomach solid and food fed and the observed per cent recovery in the laboratory (Table 3), A. fulvescens ate more food in the river (5.17 g) than in the laboratory (2.33 g).Calculation of Fulton's condition factor using the collected total length (Figure 1) and wet mass of the sampled lake sturgeon indicated significantly higher condition of wild lake sturgeon caught from the Tennessee River (0.402 ± 0.03) compared to A. fulvescens measured in the lab (0.303 ± 0.01) (p = 0.0018). Previously reported condition factors for this species at the same approximate size (0.216–0.349 m) are similar (0.36–0.41) and increased in larger size classes (i.e. 0.40–0.62 for TL > 0.35 m) (Barth & Anderson, 2015). This suggests that the significant difference in condition seen between the lab and river groups in our study may be mainly related to the larger size of the fish captured from the river, rather than nutritional status or other factors. However, sizes at age and condition of A. fulvescens caught from the Winnipeg River were highly variable (Barth & Anderson, 2015), with most of this variation attributed to the river segment, suggesting that variability in factors such as flow regime, temperature, substrate type and available diet likely influence growth rate. This highlights the need to sample fish from multiple locations within a given river system to gain a complete understanding of condition and growth of A. fulvescens in the wild.The post‐feeding time of gastric lavage had no impact on the residual stomach lipid contents over 12 h post‐feeding time. However, there was a significant reduction in protein concentration (mg/dL) at 2, 4, 6 and 12 h in stomach content (Figure 2A). Glucose concentrations (mg/dL) in both stomach and colonic contents at different time points were also significantly different (Figure 2E,F). That means A. fulvescens absorb most of the protein in the first 2 h of digestion, although the absorption rate may be variable depending on the total protein present in the ingested feed (Figure 3A). Lipid concentration (mg/dL) showed no significant difference among different time points.Efficiency of gastric lavage and colonic flushingIn this study, the efficiency of an established gastric lavage technique was determined and then utilized to assess the nutritional status of wild A. fulvescens. The gastric lavage or stomach flushing technique has been previously demonstrated to be a safe, non‐lethal method for removing food items from the stomach of fish, including Acipenser species (Brosse et al., 2002; Haley, 1998; Hyslop, 1980). In the present study, no mortality was observed during gastric lavage of A. fulvescens and for 28 days after gastric lavage was done in the laboratory; although any long‐term mortality in the Tennessee River was not able to be tracked, it is reasonable to assume similar survival rates for wild fish undergoing the procedure. This result confirms similar outcomes as previous studies regarding the safe use of the gastric lavage technique for A. fulvescens, despite having been developed for other sturgeon species (Brosse et al., 2002; Haley 1998). The gastric lavage system was easily operated by one person. The gastric lavage technique used in this study is efficient and safe for small‐sized sturgeon; however, it is time‐consuming for large fish which is associated with increased stress and delayed release of wild A. fulvescens due to the required increase in monitoring during procedure recovery time. Collection efficiency is higher in smaller fish than large fish, as smaller fish are sedated and recovered from sedation more readily than larger animals.According to Brosse et al. (2002), the recovery rate of stomach content decreases after 2 h of feeding, and thus, the procedure will only recover the most recently ingested food. We observed this increase in recovery at 4 h in captive juvenile A. fulvescens but continued to recover stomach content up to 12 h post‐feeding. In addition, stomach content was recovered from nearly all wild‐caught A. fulvescens following an overnight line set, with fish caught potentially up to 14 h prior to sampling (Table 4). Another study found that the average food recovery rate for all food items was 74.9% by number and 73.7% by weight from juvenile Pallid Sturgeon (Scaphirhynchus albus) (Wanner, 2006); in comparison the highest recovery from A. fulvescens in the present study was 75.81% ± 14.09% stomach solid by weight at 4 h post‐feeding, with most other time points substantially lower (Table 3). The recovery rate of stomach contents may be higher in A. fulvescens due to better efficiency of the gastric lavage technique.Proximate composition analysisThe knowledge of nutrient requirements and physiological systems is required to develop a successful feeding programme for sturgeon reared in hatcheries. To achieve efficient fish growth and survival, information regarding species‐specific digestive physiology, gastrointestinal anatomy and dietary preference are essential (Klasing, 2005). The results of the present study demonstrated no significant difference in protein concentration of gut content between laboratory fish and wild fish. The optimal dietary protein requirement does not differ much in different sturgeon to gain maximum ideal body weight (IBW), ranging from 37% to 45% across several species of Acipenseridae (Hung, 2017). The reason for the slight differences between species include different methodologies used, different initial weights, sources of dietary protein, other nutritional components, experimental conditions and statistical methods used among the studies, rather than true species‐specific differences (Hung, 2017). In the current study, the juvenile A. fulvescens were fed Rangen and krill in the laboratory. To our knowledge, the only reported diet for this species in the Tennessee river consists almost entirely of larval Chironomidae, mostly Chironomus spp., with Coelotanypus, Ablabesmyia, Cryptochironomus, and Procladius spp. to a lesser extent (Amacker & Alford, 2017). Although the stomach's food content was semi‐digested, the wild diet has a similar amount of protein concentration (average 40.00 ± 6.57 mg/dL) as the provided laboratory diet (average 46.17 ± 7.32 mg/dL), which is within the range of protein requirements of other sturgeon species. Fish sampled in both the laboratory and the Tennessee River absorbed the same amount of protein. So, it is likely that A. fulvescens in the Tennessee River are able to acquire the IBW by feeding the amounts within the measured range of this study (40.00–46.17 mg/dL).There was significantly higher lipid content in both stomach (average 40.11 ± 7.71 mg/dL) and colonic solids (35.59 ± 7.29 mg/dL) of fish in the laboratory than in the wild fish stomach (18.26 ± 3.47 mg/dL) and colonic contents (22.00 ± 5.54) (Figure 2). There might be two possible reasons: (1) The diet available in the wild habitat may contain less lipid than the laboratory diet or (2) the wild fish may be able to digest and absorb lipid faster than the laboratory‐reared A. fulvescens. Related to the first possibility, it could also be that lipids are present but less accessible due to the need to first digest protective chitinous or calcareous shells of ingested natural feeds. The level of dietary lipid is known to affect specific growth rates. In a study by Guo et al. (2011), when the dietary lipid level was increased from 4% to 19% with 37% crude protein and fed to hybrid sturgeon, it increased the specific growth rate. Increasing dietary lipid levels did not affect whole‐body protein level, but whole‐body lipid increased significantly with increasing dietary lipid levels. Hybrid sturgeon fed the diet with the highest lipid (19%) also had the highest whole‐body lipid content (Guo et al., 2011). According to Zheng et al. (2010), dietary lipid also affects fish survival. A study on the effect of dietary lipid on survival and growth of Dark Barbel Catfish (Pelteobagrus vachelli) showed increased dietary lipid level from 58 to 151 g/kg and increased survival from 11.6% ± 4.3% to 32.2% ± 3.8%. However, when the amount of lipid increased to 199 g/kg, the percentage of survival decreased to 17.7% ± 4.4%, which was not significantly different from the survival at 58 g/kg (Zheng et al., 2010). Given that passage rate appears to be similar in the laboratory and wild‐caught groups in the present study, it seems likely that the observed difference is attributable to inherent dietary composition differences. If so, the difference in lipid concentration may affect both the growth and survival of A. fulvescens in their native habitat during reintroduction. Thus, the scope for increasing survival by modifying the diet provided prior to stocking may be limited if available lipid‐rich dietary resources are not available in the natural system. Further research on stress and survival outcomes due to the dietary shift experienced at stocking would help determine the best recommendations for the most beneficial stocking programme modifications. Additionally, it is possible that the habitat in the area of collection was not suitable resulting in the observed low nutritional state. To our knowledge, diet utilization by A. fulvescens in the Tennessee River has been previously reported for only one reservoir (Ft. Loudoun, Amacker & Alford, 2017), but the available foods may differ between reservoirs, and the assumption that bloodworms (Table 3) are a reasonable proxy for the composition of the natural diet in Watt's Barr reservoir has not been verified. Thus, follow‐up studies of other locations with populations of A. fulvescens are needed to determine whether this is a site‐limited effect or generally true of natural A. fulvescens habitats. Rearing fish in a hatchery using diet with similar proximate composition as the wild diet may reduce the stress of stocking in the wild habitat. However, providing a similar diet as the wild diet would be predicted to provide similar growth and survival rates.The glucose concentration is low in both laboratory‐reared and wild fish, and there is no significant difference in glucose concentration of stomach content of A. fulvescens in the lab (2.33 ± 0.64 mg/dL) versus wild animals (3.57 ± 0.86 mg/dL), whereas the glucose content in colonic content of lab fish (1.34 ± 0.23 mg/dL) is significantly lower than in wild fish (4.19 ± 0.56 mg/dL). Dietary glucose has significant effects on IBW. It has been previously found that Acipenser transmontanus fed a diet without d‐glucose had a significantly lower percentage of IBW than the fish fed with 21% or more d‐glucose (Hung, 2017). One study on glycosuric responses by A. transmontanus indicates that plasma glucose concentration increased at 1‐h post‐feeding time among the seven different groups incubated with different carbohydrates (Deng et al., 2001). Although complex carbohydrates were not measured in this study, we found a significant reduction in glucose concentration in stomach contents from 0‐ to 2‐h post‐feeding time (Figure 2E), indicating that the readily available glucose from food content was absorbed into the blood plasma rapidly, within 2 h after feeding. Additionally, our results indicated significant retention of glucose in colonic contents of wild fish (Figure 3B). A likely source of complex carbohydrates in wild diets is the incidental ingestion of benthic plant matter, which is an interesting area for future research. According to Kaushik et al. (1989), Acipenser baerii did not use the complex carbohydrates such as crude starch well when they were fed with diets containing 38% of crude starch, gelatinized starch, extruded starch or 46% of extruded corn. Despite the overall small portion of the diet represented by glucose, the form in which it is present in the diet could still have substantial outcomes for carbohydrate absorption rates, and the success of this species in the wild.CONCLUSIONSGastric lavage and colonic flushing techniques have been proven efficient in both laboratory and wild‐caught A. fulvescens. Neither mortalities nor growth reduction was observed after collecting stomach content using these techniques in laboratory fish, although mortality was not tracked in the wild habitat. In the laboratory, no stomach content was collected after 12 h. The total amount of mass‐specific colonic content was more than the stomach content in the field due to prolonged post‐feeding time (14 h) before collecting contents. There was no significant difference in protein of both stomach and colonic contents between wild‐caught and laboratory‐held fish; however, the concentration of lipid was significantly higher in stomach and colonic content of laboratory A. fulvescens, and glucose was higher in the colonic contents of wild‐caught fish. Future studies on access to and use of energy‐dense foods in the river may provide information to improve survival of A. fulvescens in support of their reintroduction and conservation.AUTHOR CONTRIBUTIONSJanet Genz conceptualized the study and methodology and oversaw the project administration, including funding and resource acquisition. Formal analysis was performed by Afroza Naznin and Janet Genz. Afroza Naznin was responsible for method validation and led the investigation, supervised by Janet Genz. Afroza Naznin prepared the original draft and visualizations, with review and editing by both.ACKNOWLEDGEMENTSWe are grateful to Carlos Echevarria and Brian Hickson from the US Fish and Wildlife Service Warm Springs National Fish Hatchery for consultation on experimental design, providing lake sturgeon to study in the laboratory, providing data regarding survival rates of stocked A. fulvescens, and for field assistance. Tennessee Wildlife Resources Agency officials helped us by providing boat time and capturing lake sturgeon. We are thankful to the graduate and undergraduate students who helped us to take care of lake sturgeon in the University of West Georgia Aquatics Research Laboratory.CONFLICT OF INTEREST STATEMENTThe authors have no conflict of interests to declare.DATA AVAILABILITY STATEMENTThe data that support the findings of this study are available from the corresponding author upon reasonable request.ETHICS STATEMENTThe care and use of experimental animals complied with the animal welfare laws, guidelines and policies as outlined by the US National Research Council's Guide for the Care and Use of Laboratory Animals, the US Public Health Service's Policy on Humane Care and Use of Laboratory Animals and Guide for the Care and Use of Laboratory Animals. Local site use of experimental animals was approved by the University of West Georgia Institutional Animal Care & Use Committee (protocol #1201).REFERENCESAloisi, D., Gordon, R.R., Starzl, N.J., Walker, J.L. & Brady, T.R. 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(2015) Food transit time and site of absorption of nutrients in Gulf of Mexico Sturgeon. North American Journal of Aquaculture, 77(3), 275–280.Walker, D.J. (2017) Habitat and population assessments of the Lake sturgeon (Acipenser fulvescens) reintroduced to the upper Tennessee River. Doctoral thesis, University of Tennessee, US. https://trace.tennessee.edu/utk_graddiss/4824/Wanner, G.A. (2006) Evaluation of a gastric lavage method on juvenile Pallid sturgeon. North American Journal of Fisheries Management, 26(3), 587–591.Zheng, K., Zhu, X., Han, D., Yang, Y., Lei, W. & Xie, S. (2010) Effects of dietary lipid levels on growth, survival and lipid metabolism during early ontogeny of Pelteobagrus vachelli larvae. Aquaculture, 299(1–4), 121–127. http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Aquaculture Fish and Fisheries Wiley

Composition analysis of gastrointestinal contents of laboratory‐reared and wild lake sturgeon (Acipenser fulvescens)

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Abstract

INTRODUCTIONConservation history of Acipenser fulvescensSturgeon belong to the family Acipenseridae, one of the most widespread families of fishes worldwide, inhabiting estuaries, rivers, near‐shore oceanic environments and inland seas of the Northern hemisphere (Pikitch et al., 2005). There are 25 extant Acipenserid species (Shen et al., 2020), including the species this study focuses on, Acipenser fulvescens (Rafinesque, 1817). This species evolved in the Jurassic period 250 million years ago and is considered a “living fossil” (Birstein, 1993; Harkness & Daymond, 1961) but suffered substantial population declines due to anthropogenic activities like habitat degradation, habitat fragmentation, and overharvesting that were common at the start of the 20th century (Pollock et al., 2015). They are primarily found in the Great Lakes and Mississippi River drainage (Ferguson & Duckworth, 1997), but they are also native to the Coosa River basin which crosses north central Georgia, southeastern Tennessee and northeast Alabama.Around 1960, the original population of A. fulvescens in the Coosa, Tennessee and Cumberland Rivers became extirpated by overharvesting due to increased demand for caviar and degraded water quality related to urbanization and intensive dam construction throughout the southeastern United States (Walker, 2017). To restore A. fulvescens to those rivers, Georgia Department of Natural Resources Wildlife Resources Division (GA‐DNR) initiated a long‐term stocking programme in the Coosa and Tennessee Rivers with the goal of re‐establishing the population of this native sport fish to Georgia and address the conservation of Georgia's rare species. Juvenile A. fulvescens were first stocked by GA‐DNR in the Coosa River system from 2002 to 2005, releasing 45,091 fingerlings during this period (Bezold & Peterson, 2008). After 4 years, an assessment study showed an increase in abundance of juveniles to an overall population estimate of n = 789 (690–889, 95% confidence interval). Each cohort's survival rate ranged from 1% to 14%, depending on the size of fish stock in each year (Bezold & Peterson, 2008), and the survival rate through 2019 is estimated to be 1% overall for the Coosa River, and 2.5% for the Tennessee River. Recovery is expected to be slow due to the lengthy juvenile stage. Sexual maturity as indicated by first spawning occurs relatively late, typically at age 12–15 for males (Bruch, 1999; Bruch & Binkowski, 2002; Peterson et al., 2007), and even later for females. One advantage for stocking success is that growth rate is faster in juveniles (Peterson et al., 2007). A similar restoration programme in Lake Oneida, New York resulted in a growth rate of 145 mm per year for A. fulvescens aged 1–5 years (Jackson et al., 2002). Despite the slow rate of maturation and long‐life span of this species, this evidence suggests that traditional stocking programmes can be utilized to support and re‐establish self‐sustaining A. fulvescens populations. The ongoing efforts in the southeastern United States have been mostly successful reintroducing a stable population, but the aspect with greatest scope for improvement is in increasing stocking survival by reducing the impact of stocking stressors such as transitions in feed.Feeding of Acipenser fulvescensFeeding by A. fulvescens is characterized as generalist and opportunistic, with larvae subsisting on zooplankton (Muir et al., 2000), and after that they consume a wide variety of benthic organisms (Klassen & Peake, 2007; Nilo et al., 2006). The feeding habits of A. fulvescens vary seasonally, and it also depends on the availability of different foods in different seasons (Barth et al., 2013). Their primary foods include amphipods, oligochaetes, ephemeropteran nymphs, trichopteran larvae, mollusks and fish eggs (Nilo et al., 2006). In the Tennessee River, juveniles have been shown to feed selectively on larval Chironomids which vary seasonally in availability (Amacker & Alford, 2017).In the hatchery, chopped or whole bloodworms (Chironomus larvae), brine shrimp (Artemia) nauplii or other raw foods are commonly fed to A. fulvescens at the larval and adult stages (Table 1) (Aloisi et al., 2006; Klassen & Peake, 2007, 2008). Fishes fed with live feeds typically have a higher growth rate and lower mortality rate than those provided formulated food (Bauman et al., 2016). Still, live foods can have many disadvantages like high cost, irregular availability, and uncertainty about nutritional quality. According to Dvorak (2009), there is a risk of transmitting an unknown disease from hatchery‐reared fish to the environment if live feeds are fed to fishes. Formulated feed is generally more cost‐effective, is easier to manage and can be developed with stable and enriched nutrients (Hamre et al., 2013). In one study, an experimental diet with 3%–6% krill or fish protein content significantly improved growth of juvenile A. fulvescens and furthermore indicated that stimulation of feeding activity in this species shifted over the course of juvenile growth (from 2 to 280 g) (Moreau & Dabrowski, 1996). Juvenile A. fulvescens can be fed formulated diets (soft‐moist or dry) without impairing their survival and growth performance (Lee et al., 2018), but the formulated diets available for sturgeon almost certainly differ in their nutritional value from the wild diet (Table 1). At stocking, the transitions from one diet in captivity to a different food source in the wild may create additional stocking stress and act as a barrier to high post‐stocking survival rates of captively reared juveniles. Therefore, it is crucial to know about the composition of both captive and wild diets of juvenile A. fulvescens to gain an understanding of nutritional impacts occurring at stocking in order to optimize programmes that aim to help restore severely depleted or extirpated populations.1TABLEComposition of provided laboratory feeds per the supplier's guaranteed analysisDietSoft‐moist pelleted feedaFrozen krillbFrozen bloodwormsbProtein (%)45.06.53.5Fat (%)19.03.40.3Fibre (%)<2.00.70.7Ash (%)<9.01.31.6Moisture (%)25.088.093.5Note: Bloodworms approximate primary foodstuffs consumed by lake sturgeon in the Tennessee river (Amacker & Alford, 2017).aRangen Brand.bSan Francisco Bay Brand.Study objectivesGrowth of fish and feed utilization are influenced by the balanced nutritional composition of fish feed. An adequate supply of nutrients, both in terms of quantity and quality, should be met for good growth, survival and reproduction of fish and other aquatic animals (Hasan, 2001). During the transition from endogenous to exogenous feeding and for several weeks after the onset of exogenous feeding, larval fish in aquaculture typically exhibit periods of low growth and high mortality (Kamali et al., 2006). Similarly, A. fulvescens experience a shift in diet composition and availability during stocking to the river, and this may be associated with reduced survival. Therefore, it is important to evaluate and assess the proximate composition of food from both laboratory and wild habitats to allow for comparison of the nutritional value in these two systems and estimate the scale of the impact this dietary shift has post‐stocking. It is also required to know about the physiological system, dietary preference, digestive physiology and gastrointestinal anatomy to develop a successful feeding programme for A. fulvescens (Hofer et al., 1982). This study assessed the nutrient uptake from the diet of laboratory‐reared and wild A. fulvescens and how the hatchery rearing environment and riverine habitat differ from each other in available nutrition. In the present study, the stomach and colonic contents of wild A. fulvescens needed to be collected to assess the proximate composition of the diet. However, it is not feasible to sacrifice A. fulvescens to know the content of this fish's food due to its reduced population status. A safe and efficient technique was also needed to ensure a quick recovery by the wild‐caught fish after collecting contents from the gastrointestinal tract. Therefore, established techniques for digesta collection were first validated using controlled laboratory trials and then applied to samples collected from wild fish. The collected samples were then analysed for proximate composition in order to provide insight to nutrient processing in hatchery‐raised A. fulvescens fed a formulated diet, compared to juveniles subsisting on the natural diet available in the wild. We first validated gastric lavage and colonic flushing in the lab to support the collection of stomach and faecal contents from wild fish and then compared the proximate composition of stomach and colonic contents between the laboratory and wild environment. This comparison will support the conservation of this protected species, helping to re‐establish the population throughout its endemic range.MATERIALS AND METHODSExperimental animalsFour 150‐gal freshwater tanks were set up with water quality and physical conditions according to standard parameters for A. fulvescens (21°C, pH 7.18 ± 0.02, conductivity 470 ± 3.21 μS, dissolved oxygen held at 100% air saturation by continuous aeration). Nineteen 2.5 year‐old A. fulvescens individuals, ranging from 0.39 to 0.56 m total length (Figure 1) and weighing 0.34 ± 0.04 kg, were received from the US Fish & Wildlife Service Warm Springs National Fish Hatchery, brought to the University of West Georgia and evenly distributed among the prepared tanks. Fish were fed 2.4 mm diameter Rangen pellets and krill (Table 1) to satiation daily at 8 AM, following the standard diet and feeding procedures of the rearing hatchery. Leftover food was siphoned out after feeding activity stopped (approximately 45 min) to maintain water quality. Fish were acclimated to laboratory conditions for 1 month prior to the start of experiments, and water quality was monitored by measuring the concentrations of nitrate, nitrite, ammonia, pH, hardness and conductivity every day for the first 15 days after introduction of fish into the laboratory environment, and after that water quality parameters were checked once every 3 days, with a daily water exchange of 20–30 gal of water in each tank.1FIGUREComparison of total length (mm) of juvenile Acipenser fulvescens utilized for validation of gastric lavage and colonic flushing protocols versus wild A. fulvescens captured from the Tennessee River.Techniques for collection of gastrointestinal contentsGastric lavage is a non‐lethal method for examining fish stomach content from live fish, where stomach contents are flushed out with an apparatus inserted into the fish's pharynx. The gastric lavage technique has been tested in several sturgeon species. However, most of these studies have analysed the stomach contents for identifying prey items or determining mortality rates of the procedure itself, rather than determination of dietary nutritional profile (Haley, 1998; Wanner, 2006). Colonic flushing is a non‐lethal method for examining faeces in the intestinal tract. Deionized (DI) water is pulsed into the colon through the anus using a syringe attached with a smaller diameter tube. The tube is inserted gently into the colon and creates short pulses to expel all the faeces. This process continues until the expelled water is clear. Colonic flushing is less time‐consuming (15–30 s) and safer for A. fulvescens than the gastric lavage technique (Amacker, 2016).Laboratory validation of gastric lavage and colonic flushingAfter acclimatizing fish to the laboratory environment, we performed non‐lethal gastric lavage by following the protocol as described by Haley (1998). Before starting gastric lavage, total weight and total length of each fish were recorded, and then gastric lavage and colonic flushing were performed at six different time points (2, 4, 6, 12, 24 and 32 h after feeding) to determine the required digestion time, using 18 randomly selected fish (n = 3 fish per time point) in two trials. In the first trial, 12 fish were used to perform gastric lavage at 2, 4, 6 and 12 h post‐feeding. In the second trial, gastric lavage was done at 4 and 12 h post‐feeding on six fish, and the same fish were used for gastric lavage at 24 and 32 h post‐feeding after 1 week. Stomach content at 0 h post‐feeding was represented by analysis of the offered feed. Colonic flushing was not performed at 0 h post‐feeding as the colon was assumed to be empty at this time point. Fish were fasted the day before performing gastric lavage and fed Rangen and krill at the start of each trial. Table 1 shows the composition of these feed as given by the manufacturer's guaranteed analysis. Ingestion of feed by size‐matched fish was assumed to be evenly distributed within the tank. When the fish's feeding activity stopped, all leftover food was siphoned from the tank. The water was filtered with a very fine mesh net and the leftover food was weighed to determine the total feed ingested. For the gastric lavage protocol, each fish was anesthetized in a 5‐gal bucket containing 30 mL of tricaine methanesulfonate (MS‐222; 100 mg/L) buffered with sodium bicarbonate (200 mg/L) combined with 20 L of water, for a final immersion concentration of 0.15 mg/L MS‐222. Typically, fish were sedated after approximately 4 min. Once sedated, the fish was placed ventral side up at a 45° angle on a gastric lavage stand where flexible tubing attached to a 60‐cm3 syringe fitted with a 16‐ga needle was inserted into the stomach through the pharynx. A different diameter tube was used for different‐sized fish (Table 2), and the large diameter tubing omitted the needle fitting and was instead connected directly to the syringe. Pulsing DI water at the cardiac valve stimulated relaxation and allowed entry of the tube into the stomach. Water was pulsed by syringe into the stomach until it was firm. Then, a gentle massage of the ventral surface of the sturgeon was done to help release water and stomach contents into a pre‐weighed sample collection bottle. This procedure was continued until only clean water came out from the stomach. Haley (1998) suggested pulsing DI water for 3 min and repeated it three times for each animal. However, in this study, the timing to complete gastric lavage depended on fish size and the amount of food they had in their stomach but never exceeded 5 min. After gastric lavage, the fish was kept in a separate bucket filled with aerated fresh water until it recovered from the stress of the procedure and resumed regular activity. For most fish, some stomach content came out during the recovery period, and this was collected using a disposable pipette.2TABLEDiameter of gastric lavage tube according to fish size (Acipenser fulvescens)Fish total length (m)Outer diameter (OD) of lavage tube (mm)Inner diameter (ID) of lavage tube (mm)0.15–0.302.421.660.31–0.454.003.150.46–1.05.154.00Colonic flushing is a non‐lethal process to examine the faeces of the intestinal tract. This process used a 60‐cm3 syringe fitted with appropriate diameter tubing (Table 2) filled with DI water and inserted the tube end gently 30–50 mm into the colon through the anus. Faecal content was collected in a bottle using the same process as described above for gastric lavage and measured for total weight. Then, the solid and aqueous parts of both stomach and colonic contents were separated using a very fine mesh filter and centrifuging the samples. The weight of both aqueous and solid parts was measured, and the samples were subsequently stored at −20°C.Stomach and colonic contents collection from wild Acipenser fulvescensStomach and colonic contents were collected from A. fulvescens caught from the upper Tennessee River over 2 days in December 2019. Trotlines were used to catch the fish from the Watts Bar Reservoir in the drainage of this river (35.6206°N, 84.7832°W). The water temperature was 10.9°C, and air temperature was 4°C on the first day of sample collection and on the second day water temperature and air temperature were 10.3 and 2°C, respectively. Dissolved oxygen in the river was variable between sampling times, on average 10.38 ± 0.32 mg/L. Trotlines were set in the river with buffalo and carp baits at 4:30 PM and retrieved at 7 AM the following day (14.5 h total soak time). After getting the fish on the boat, the total and fork lengths and the total weight of each fish were measured. Once the trotlines were removed from the water, fish were transported in a live well to the riverbank where a 150‐gal tank was set up for keeping fish while collecting gut and colonic contents. Stomach and colonic contents of these different sized A. fulvescens (n = 15) were collected using the same procedures described before for laboratory fish and then released to the river. The size of tubing used for the gastric lavage was varied according to the fish's size, as given in Table 2. Wild fish had an average total length of 0.65 ± 0.74 m, ranging from 0.26 to 1.08 m (Figure 1). A tube of the same diameter (inner diameter 1.14 mm, outer diameter 1.57 mm) was used to collect colonic content for every fish size to provide more space for colonic content to come out. The bottles of the stomach and colonic content were transported on ice in a cooler to the University of West Georgia Aquatics Research Laboratory and stored at 4°C prior to filtration and processing and subsequently stored at −20°C for further analysis.Proximate composition analysis of stomach and colonic samplesAll stomach contents and faecal samples collected from wild and hatchery‐reared A. fulvescens were prepared and analysed according to the same protocols. The solid portion of the sample was separated by filtration using fine nylon mesh (approx. 0.3–0.5 mm pore size) followed by centrifugation at 10,000 × g for 10 min and stored at −20°C. Each solid sample was diluted by adding 2x (by weight) of the corresponding aqueous portion previously separated by filtration and then an electric tissue homogenizer was used for homogenization. Thus, both the solid and liquid fractions of the original sample were represented in the analysed sample. For the 0 h time point, thawed krill and Rangen pellets were separately diluted and homogenized. Heating of homogenate was prevented by using a short pulse (<5 s). The crude homogenate was separated into three aliquots for analysis of lipid, protein and carbohydrate. Aliquots for the water‐soluble protein and glucose fractions were centrifuged at 10,000 × g for 10 min to remove solid fragments, and the supernatant transferred to a new tube for analysis. The Folch method (Folch et al., 1957) was used for extraction of lipids from the remaining aliquot. In brief, 20× by volume (1500 μL) of 2:1 (v/v) chloroform:methanol solution was added to a 75 μL aliquot of the crude homogenate, mixed by vortexing and then kept on ice for 5 min. Then 0.2× (315 μL) 0.05% CaCl2 solution was added and centrifuged at 12,000 × g for 5 min. After centrifuging, the lower phase was separated to a new tube and used to measure the lipid content (mg/dL).Each processed sample aliquot was analysed for proximate composition using colorimetric assay kits, according to the manufacturer's protocols. Total protein (Sigma‐Aldrich, 96 Well Plate Bradford Assay Protocol, B6916), triglyceride (Cayman Chemical, No. 10010303) and glucose (Cayman Chemical, No. 10009582) were measured to give the concentration (mg/dL) of each of these macronutrients in each collected gut sample. In the Sigma protein assay protocol, a protein standard of bovine serum albumin (10–160 mg/dL) was used to make the standard curve, and 5 μL of each unknown sample was pipetted into other wells on the same plate. After initiating the reaction with Bradford reagent, the absorbance was measured at 595 nm at 25°C. For the lipid assay, 10 μL triglyceride standard of 0–200 mg/dL concentration was used for the standard curve, and 10 μL of each unknown sample was measured on a 96‐well plate, according to the kit manufacturer's instructions. The plate was incubated at 25°C for 15 min after adding 150 μL diluted enzyme buffer. Then the lipid concentration was measured by absorbance at 540 nm. For measurement of the glucose concentration, 15 μL of standard solution at 0–25 mg/dL concentrations established the standard curve. To each well, 85 μL of diluted NP‐40 assay buffer was added, and 100 μL of reconstituted enzyme mixture initiated the reaction. Absorbance was measured at 520 nm after incubation at 37°C for 10 min. Each unknown sample and standard solutions were measured in triplicate for all protocols.Statistical analysisAll data are presented as a mean ± SEM and were analysed using GraphPad Prism software (version 6.07). Multiple t‐tests were used to compare concentration of protein, lipid and glucose between the treatment groups. To compare the concentrations of protein, lipid and glucose among different time points, one‐way ANOVA was used, followed by Tukey's multiple comparisons post hoc test to examine pairwise comparisons. Values of p < 0.05 were considered significant, represented by an asterisk (*).RESULTSEfficiency of gastric lavage and colonic flushingGastric lavage and colonic flushing were used to collect stomach and colonic contents in both laboratory‐reared and wild A. fulvescens. The highest average percentage of ingested feed, 75.82% ± 14.09%, was recovered from the stomach at 4 h post‐feeding, with substantially lower amounts recovered at 2 h (36.67% ± 5.09%), 6 h (17.80 ± 4.01) and at 12 h (32.90% ± 17.93%) in the laboratory. No food was recovered after 12 h (Table 3). The amount of recovered colonic content increased with increasing time, and the amount per fish body mass (g/kg) was highest at 32 h post‐feeding (Table 3).3TABLEAmount of stomach and colonic content recovered from Acipenser fulvescens using gastric lavage and colonic flushing at 2–32 h post‐feeding in the laboratory (n = 3–6 per time point) as a percent of feed ingested (%), and relative to fish mass (g/kg)Post‐feeding time (h)Recovered stomach content (%)Stomach content (g/kg)Colonic content (g/kg)236.67 ± 5.090.29 ± 0.050.63 ± 0.29475.82 ± 14.090.96 ± 0.431.00 ± 0.40617.80 ± 4.010.18 ± 0.041.16 ± 0.401232.90 ± 17.930.32 ± 0.141.26 ± 0.3224003.01 ± 0.7932005.40 ± 1.30Average27.20 ± 11.620.29 ± 0.142.08 ± 0.75Note: Fish were fed a total of 0.33 g Rangen soft‐moist pellets and 2.33 g krill per fish.Following the protocol of gastric lavage and colonic flushing used in the laboratory, stomach and colonic contents were collected from A. fulvescens captured from the Tennessee River. We used a total of 15 fish for gastric lavage and colonic flushing in the wild habitat. One fish was not used for gastric lavage among those fish, and another fish was not used for colonic flushing because those fish were not in good enough condition to remain out of the water for the time required to perform both procedures. The amount of food ingested by A. fulvescens in the river was estimated based on the amount of recovered stomach solids, using digestion rates observed in the lab fish. In the lab, the average recovered stomach solid over 12 h was 1.23 g, and the amount of krill ingested per fish was 2.33 g (Table 3), indicating a recoverable proportion of 1.89, which was multiplied by the mean weight of stomach solids recovered from wild fish (2.72 g, Table 4) resulting in an estimated 5.15 g of food ingested by wild fish prior to capture. The average stomach solid recovery rate adjusted for fish mass (1.41 ± 0.51 g/kg) was lower than the colonic solid recovery rate (5.85 ± 0.94 g/kg) (Table 4). In wild fish, variation between individuals was greater, but on average, the amounts of solid content collected from the stomach and distal intestine were both roughly threefold greater compared to the A. fulvescens sampled in the lab (Table 3).4TABLEAmount of stomach and colonic content collected from Acipenser fulvescens (n = 15) by gastric lavage or colonic flushing at Tennessee RiverFish mass (kg)Stomach solid (g)Stomach solid recovered (g/kg)Colonic solid (g)Colonic solid recovered (g/kg)1.590.320.197.614.791.912.211.157.003.672.042.901.429.834.801.092.332.109.008.231.130.300.267.466.531.9513.807.08N/AN/A1.870.310.161.520.801.360.620.4416.9012.421.465.703.919.566.512.340.810.3417.127.281.630.530.318.124.901.630.800.499.655.881.891.440.7423.2112.295.266.211.1842.548.086.44N/AN/A9.931.54Average2.24 ± 0.392.73 ± 0.991.41 ± 0.5112.82 ± 2.695.85 ± 0.94Composition of stomach contents at different time pointsA total of 18 fish were used to evaluate gastric lavage efficiency in the laboratory at 2, 4, 6, 12, 24 and 32 h after feeding. The proximate composition was analysed individually at each sampling time. There was a significant difference in protein concentration in stomach (p = 0.0074, Figure 2A) over time but not in the colonic content (p = 0.3853, Figure 2B) over the six different time points examined in laboratory‐held A. fulvescens. Tukey's multiple comparison test identified that protein content was significantly lower than the provided feed (0 h) at every time point for which stomach contents were recovered (2, 4, 6 and 12 h). Lipid concentration (mg/dL) in recovered stomach and colonic contents was also assessed at the same post‐feeding time points. There was no significant difference in lipid concentration in stomach (p = 0.3174, Figure 2C) or colonic content (p = 0.1057, Figure 2D) among the measured time points in laboratory A. fulvescens. Glucose concentration in both stomach (p = 0.0240, Figure 2E) and colonic (p = 0.0480, Figure 2F) content was significantly different over the observed time frame. Interestingly, Tukey's multiple comparison test did not identify any significant differences among different time points in colonic glucose concentration (mg/dL), indicating that only the overall difference was significant for these samples. However, in stomach content, it showed a significant difference between 0 h compared to all subsequent time points (2, 4, 6 and 12 h).2FIGUREProtein, lipid and glucose concentration (mg/dL) in stomach (A, C, E) and colonic (B, D, F) contents from laboratory acclimated Acipenser fulvescens at 0–32 h post‐feeding. Stomach values at 0 h are the measured concentrations in the offered feed as determined by analysis identical to methods used for collected digesta. Colonic samples were not collected at 0 h. One‐way ANOVA indicated a significant difference in protein concentration in stomach content (p = 0.0074, A) but not in colonic content (p = 0.3853, B). Lipid concentrations (mg/dL) in both stomach (p = 0.3174, C) and colonic (p = 0.1057, D) contents at different post‐feeding time points were not significantly different. Glucose concentration (mg/dL) in stomach (p = 0.0240, E) and colonic (p = 0.0480, F) contents at different post‐feeding time‐points shows significant differences. Post hoc differences from 0 h, as determined by a pairwise Tukey's post hoc test, are indicated by “*”.Proximate composition of field‐collected stomach and colonic contentSeparated solid content from stomach and colon were used for proximate composition analysis by colorimetric assay protocol. Protein concentrations in stomach contents between laboratory fishes and wild fish were not significantly different (p = 0.2104) (Figure 3A), whereas there was a significant difference (p = 0.0187) in lipid concentration of stomach contents, with A. fulvescens fed the laboratory diet exhibiting substantially greater lipid content than wild‐collected fish. The stomach contents of fish from both habitats showed no difference (p = 0.6835) in glucose concentration (Figure 3A). A slightly different pattern was observed for colonic solids, with no differences between sampling groups in protein or lipid content, but glucose was significantly higher in wild fish (Figure 3B). Glucose was by far the smallest fraction contributing to the proximate composition of stomach and colonic contents of A. fulvescens in both the laboratory and wild habitats.3FIGUREComparison of protein, lipid and glucose concentrations (mg/dL) in the stomach solids (A) and colonic solids (B) of laboratory‐reared and wild Acipenser fulvescens. Asterisk symbol “*” indicates significant difference (p < 0.05) in concentrations between wild and laboratory‐reared fish as determined by multiple t‐tests, for protein (p = 0.2104), lipid (p = 0.0187) and glucose (p = 0.6835) of stomach contents (A) and protein (p = 0.6198), lipid (p = 0.1483) and glucose (p = 2.52e – 006) of faeces (B).DISCUSSIONFood transit time in alimentary canalThe amount of colonic content increased with time after feeding the fish in the laboratory. According to Venero et al. (2015), defecation is started between 8 and 16 h after feeding in the Gulf Sturgeon (Acipenser oxyrinchus desotoi), but in the present study faecal content was collected as early as 2 h after feeding. The fish were fasted for 24 h before each round of colonic sampling. As the total food transit time for A. fulvescens has been previously reported to be 32 h at 21°C (Venero et al., 2015), the faecal content collected at 2 h post‐feeding must have been from the day before colonic flushing was performed, that is, the residual of food ingested prior to the fasting period.The colonic flushing technique was efficient for all fish between 0.26 and 1.08 m, and larger amounts of faecal content on a per mass basis were collected from most of the wild fish than from the laboratory fish. Comparatively, the amount of faecal content was more than the amount of stomach content in wild A. fulvescens. The reason may be attributed to the amount of time the fish was on the hook of the trotline. The trotlines were in the river for approximately 14 h; if the fish was hooked at the beginning of the soak time, then it was not able to eat anything during this period. Thus, all the stomach contents will be digested, and the colonic content will increase, but as our laboratory experiments confirm, would likely still be present in the alimentary tract. Aside from consumption of the catch bait, the timing of the most recent feeding event for wild A. fulvescens cannot be known. The average amount of stomach solid recovered (1.41 ± 0.51 g/kg) from wild fish was higher than the laboratory fish (0.29 ± 0.14 g/kg); however, the weight‐specific collection rate for wild fish was quite similar to the laboratory‐based validation trial, indicating that recovered stomach contents were equally representative of the wild diet as in the laboratory validation trial, and was being digested at a comparable rate despite the 10°C difference between water temperatures in the laboratory and the river. As estimated by back calculating from the recovered amount of stomach solid and food fed and the observed per cent recovery in the laboratory (Table 3), A. fulvescens ate more food in the river (5.17 g) than in the laboratory (2.33 g).Calculation of Fulton's condition factor using the collected total length (Figure 1) and wet mass of the sampled lake sturgeon indicated significantly higher condition of wild lake sturgeon caught from the Tennessee River (0.402 ± 0.03) compared to A. fulvescens measured in the lab (0.303 ± 0.01) (p = 0.0018). Previously reported condition factors for this species at the same approximate size (0.216–0.349 m) are similar (0.36–0.41) and increased in larger size classes (i.e. 0.40–0.62 for TL > 0.35 m) (Barth & Anderson, 2015). This suggests that the significant difference in condition seen between the lab and river groups in our study may be mainly related to the larger size of the fish captured from the river, rather than nutritional status or other factors. However, sizes at age and condition of A. fulvescens caught from the Winnipeg River were highly variable (Barth & Anderson, 2015), with most of this variation attributed to the river segment, suggesting that variability in factors such as flow regime, temperature, substrate type and available diet likely influence growth rate. This highlights the need to sample fish from multiple locations within a given river system to gain a complete understanding of condition and growth of A. fulvescens in the wild.The post‐feeding time of gastric lavage had no impact on the residual stomach lipid contents over 12 h post‐feeding time. However, there was a significant reduction in protein concentration (mg/dL) at 2, 4, 6 and 12 h in stomach content (Figure 2A). Glucose concentrations (mg/dL) in both stomach and colonic contents at different time points were also significantly different (Figure 2E,F). That means A. fulvescens absorb most of the protein in the first 2 h of digestion, although the absorption rate may be variable depending on the total protein present in the ingested feed (Figure 3A). Lipid concentration (mg/dL) showed no significant difference among different time points.Efficiency of gastric lavage and colonic flushingIn this study, the efficiency of an established gastric lavage technique was determined and then utilized to assess the nutritional status of wild A. fulvescens. The gastric lavage or stomach flushing technique has been previously demonstrated to be a safe, non‐lethal method for removing food items from the stomach of fish, including Acipenser species (Brosse et al., 2002; Haley, 1998; Hyslop, 1980). In the present study, no mortality was observed during gastric lavage of A. fulvescens and for 28 days after gastric lavage was done in the laboratory; although any long‐term mortality in the Tennessee River was not able to be tracked, it is reasonable to assume similar survival rates for wild fish undergoing the procedure. This result confirms similar outcomes as previous studies regarding the safe use of the gastric lavage technique for A. fulvescens, despite having been developed for other sturgeon species (Brosse et al., 2002; Haley 1998). The gastric lavage system was easily operated by one person. The gastric lavage technique used in this study is efficient and safe for small‐sized sturgeon; however, it is time‐consuming for large fish which is associated with increased stress and delayed release of wild A. fulvescens due to the required increase in monitoring during procedure recovery time. Collection efficiency is higher in smaller fish than large fish, as smaller fish are sedated and recovered from sedation more readily than larger animals.According to Brosse et al. (2002), the recovery rate of stomach content decreases after 2 h of feeding, and thus, the procedure will only recover the most recently ingested food. We observed this increase in recovery at 4 h in captive juvenile A. fulvescens but continued to recover stomach content up to 12 h post‐feeding. In addition, stomach content was recovered from nearly all wild‐caught A. fulvescens following an overnight line set, with fish caught potentially up to 14 h prior to sampling (Table 4). Another study found that the average food recovery rate for all food items was 74.9% by number and 73.7% by weight from juvenile Pallid Sturgeon (Scaphirhynchus albus) (Wanner, 2006); in comparison the highest recovery from A. fulvescens in the present study was 75.81% ± 14.09% stomach solid by weight at 4 h post‐feeding, with most other time points substantially lower (Table 3). The recovery rate of stomach contents may be higher in A. fulvescens due to better efficiency of the gastric lavage technique.Proximate composition analysisThe knowledge of nutrient requirements and physiological systems is required to develop a successful feeding programme for sturgeon reared in hatcheries. To achieve efficient fish growth and survival, information regarding species‐specific digestive physiology, gastrointestinal anatomy and dietary preference are essential (Klasing, 2005). The results of the present study demonstrated no significant difference in protein concentration of gut content between laboratory fish and wild fish. The optimal dietary protein requirement does not differ much in different sturgeon to gain maximum ideal body weight (IBW), ranging from 37% to 45% across several species of Acipenseridae (Hung, 2017). The reason for the slight differences between species include different methodologies used, different initial weights, sources of dietary protein, other nutritional components, experimental conditions and statistical methods used among the studies, rather than true species‐specific differences (Hung, 2017). In the current study, the juvenile A. fulvescens were fed Rangen and krill in the laboratory. To our knowledge, the only reported diet for this species in the Tennessee river consists almost entirely of larval Chironomidae, mostly Chironomus spp., with Coelotanypus, Ablabesmyia, Cryptochironomus, and Procladius spp. to a lesser extent (Amacker & Alford, 2017). Although the stomach's food content was semi‐digested, the wild diet has a similar amount of protein concentration (average 40.00 ± 6.57 mg/dL) as the provided laboratory diet (average 46.17 ± 7.32 mg/dL), which is within the range of protein requirements of other sturgeon species. Fish sampled in both the laboratory and the Tennessee River absorbed the same amount of protein. So, it is likely that A. fulvescens in the Tennessee River are able to acquire the IBW by feeding the amounts within the measured range of this study (40.00–46.17 mg/dL).There was significantly higher lipid content in both stomach (average 40.11 ± 7.71 mg/dL) and colonic solids (35.59 ± 7.29 mg/dL) of fish in the laboratory than in the wild fish stomach (18.26 ± 3.47 mg/dL) and colonic contents (22.00 ± 5.54) (Figure 2). There might be two possible reasons: (1) The diet available in the wild habitat may contain less lipid than the laboratory diet or (2) the wild fish may be able to digest and absorb lipid faster than the laboratory‐reared A. fulvescens. Related to the first possibility, it could also be that lipids are present but less accessible due to the need to first digest protective chitinous or calcareous shells of ingested natural feeds. The level of dietary lipid is known to affect specific growth rates. In a study by Guo et al. (2011), when the dietary lipid level was increased from 4% to 19% with 37% crude protein and fed to hybrid sturgeon, it increased the specific growth rate. Increasing dietary lipid levels did not affect whole‐body protein level, but whole‐body lipid increased significantly with increasing dietary lipid levels. Hybrid sturgeon fed the diet with the highest lipid (19%) also had the highest whole‐body lipid content (Guo et al., 2011). According to Zheng et al. (2010), dietary lipid also affects fish survival. A study on the effect of dietary lipid on survival and growth of Dark Barbel Catfish (Pelteobagrus vachelli) showed increased dietary lipid level from 58 to 151 g/kg and increased survival from 11.6% ± 4.3% to 32.2% ± 3.8%. However, when the amount of lipid increased to 199 g/kg, the percentage of survival decreased to 17.7% ± 4.4%, which was not significantly different from the survival at 58 g/kg (Zheng et al., 2010). Given that passage rate appears to be similar in the laboratory and wild‐caught groups in the present study, it seems likely that the observed difference is attributable to inherent dietary composition differences. If so, the difference in lipid concentration may affect both the growth and survival of A. fulvescens in their native habitat during reintroduction. Thus, the scope for increasing survival by modifying the diet provided prior to stocking may be limited if available lipid‐rich dietary resources are not available in the natural system. Further research on stress and survival outcomes due to the dietary shift experienced at stocking would help determine the best recommendations for the most beneficial stocking programme modifications. Additionally, it is possible that the habitat in the area of collection was not suitable resulting in the observed low nutritional state. To our knowledge, diet utilization by A. fulvescens in the Tennessee River has been previously reported for only one reservoir (Ft. Loudoun, Amacker & Alford, 2017), but the available foods may differ between reservoirs, and the assumption that bloodworms (Table 3) are a reasonable proxy for the composition of the natural diet in Watt's Barr reservoir has not been verified. Thus, follow‐up studies of other locations with populations of A. fulvescens are needed to determine whether this is a site‐limited effect or generally true of natural A. fulvescens habitats. Rearing fish in a hatchery using diet with similar proximate composition as the wild diet may reduce the stress of stocking in the wild habitat. However, providing a similar diet as the wild diet would be predicted to provide similar growth and survival rates.The glucose concentration is low in both laboratory‐reared and wild fish, and there is no significant difference in glucose concentration of stomach content of A. fulvescens in the lab (2.33 ± 0.64 mg/dL) versus wild animals (3.57 ± 0.86 mg/dL), whereas the glucose content in colonic content of lab fish (1.34 ± 0.23 mg/dL) is significantly lower than in wild fish (4.19 ± 0.56 mg/dL). Dietary glucose has significant effects on IBW. It has been previously found that Acipenser transmontanus fed a diet without d‐glucose had a significantly lower percentage of IBW than the fish fed with 21% or more d‐glucose (Hung, 2017). One study on glycosuric responses by A. transmontanus indicates that plasma glucose concentration increased at 1‐h post‐feeding time among the seven different groups incubated with different carbohydrates (Deng et al., 2001). Although complex carbohydrates were not measured in this study, we found a significant reduction in glucose concentration in stomach contents from 0‐ to 2‐h post‐feeding time (Figure 2E), indicating that the readily available glucose from food content was absorbed into the blood plasma rapidly, within 2 h after feeding. Additionally, our results indicated significant retention of glucose in colonic contents of wild fish (Figure 3B). A likely source of complex carbohydrates in wild diets is the incidental ingestion of benthic plant matter, which is an interesting area for future research. According to Kaushik et al. (1989), Acipenser baerii did not use the complex carbohydrates such as crude starch well when they were fed with diets containing 38% of crude starch, gelatinized starch, extruded starch or 46% of extruded corn. Despite the overall small portion of the diet represented by glucose, the form in which it is present in the diet could still have substantial outcomes for carbohydrate absorption rates, and the success of this species in the wild.CONCLUSIONSGastric lavage and colonic flushing techniques have been proven efficient in both laboratory and wild‐caught A. fulvescens. Neither mortalities nor growth reduction was observed after collecting stomach content using these techniques in laboratory fish, although mortality was not tracked in the wild habitat. In the laboratory, no stomach content was collected after 12 h. The total amount of mass‐specific colonic content was more than the stomach content in the field due to prolonged post‐feeding time (14 h) before collecting contents. There was no significant difference in protein of both stomach and colonic contents between wild‐caught and laboratory‐held fish; however, the concentration of lipid was significantly higher in stomach and colonic content of laboratory A. fulvescens, and glucose was higher in the colonic contents of wild‐caught fish. Future studies on access to and use of energy‐dense foods in the river may provide information to improve survival of A. fulvescens in support of their reintroduction and conservation.AUTHOR CONTRIBUTIONSJanet Genz conceptualized the study and methodology and oversaw the project administration, including funding and resource acquisition. Formal analysis was performed by Afroza Naznin and Janet Genz. Afroza Naznin was responsible for method validation and led the investigation, supervised by Janet Genz. Afroza Naznin prepared the original draft and visualizations, with review and editing by both.ACKNOWLEDGEMENTSWe are grateful to Carlos Echevarria and Brian Hickson from the US Fish and Wildlife Service Warm Springs National Fish Hatchery for consultation on experimental design, providing lake sturgeon to study in the laboratory, providing data regarding survival rates of stocked A. fulvescens, and for field assistance. Tennessee Wildlife Resources Agency officials helped us by providing boat time and capturing lake sturgeon. 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Journal

Aquaculture Fish and Fisheries – Wiley

Published: Apr 1, 2023

Keywords: conservation; feed utilization; fish nutrition; gastric lavage; sturgeon; Tennessee River

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